Nematode collection and materials examined

Naturally nematode-infested experimental fields of CIMMYT located in two different regions in the Central Anatolian Plateau of Turkey including Yozgat (39° 08ʼ N, 34° 10ʼ E; altitude, 985 m) and Haymana (39° 25ʼ 52ʼ N, 39° 29ʼ 44ʼ E; altitude, 1259 m) were sampled at crop maturity in 2013. Nematode cysts were extracted from rhizosphere and roots of nematode susceptible and resistant wheat varieties using the modified flotation decanting method [27] and handpicked from the extracted suspensions under a Leitz dissecting microscope. Cysts were stored in 1.5 mL microtubes at 4°C either in dry condition or in water for further experiments. Reference strains required for taxonomic and phylogenetic inferences were obtained from the Westerdijk Fungal Biodiversity Institute (Utrecht, The Netherlands).

Ethics statement

Both the sampling sites and the nematode species were not protected and thus no permits were required.

Fungal isolation from field collected samples

Homogenously brown and visibly emptied cysts of H. filipjevi were separated from hereafter called symptomatic cysts showing unusual discolourations or fungal colonization. Under a laminar flow hood, symptomatic cysts were surface sterilized in 0.5% sodium hypochlorite (NaOCl) for 10 min and rinsed six times with sterile deionised water (SDW). The sterilizing effect of NaOCl was evaluated. For this, individual cysts were imprinted into potato dextrose agar medium (PDA; Merck, Germany) using a sterile forceps, and immediately transferred to new PDA plates. The control plates were incubated at room temperature and regularly monitored for contaminants for four weeks to exclude not-successfully surface-sterilised cysts from further analyses. Using a sterile forceps and an insect needle, transferred cysts were separately cut open on the fresh agar medium and eggs were dispersed on the agar plate. Cyst debris, i.e. particles of the cyst wall, was discarded and nematode eggs showing symptoms of fungal infections were rolled gently on agar surface to remove potentially occurring contaminating fungal propagules or hyphae adhering to the surface of eggs. Eggs were then transferred to a sterile watch glass containing SDW. When settled, eggs were rinsed six times by removing and replacing the majority of SDW. Eggs were then surface sterilized in 0.5% NaOCl for 2 min. The disinfection solution was removed and the eggs were washed up to six times with SDW as described above.

Surface-sterilised eggs of each individual cyst were then transferred to a new agar plate using a pipette and divided in two aliquots. Eggs from one portion were individually picked up and placed on PDA amended with penicillin G (240 mg/L) and streptomycin sulphate (200 mg/L) (PDA+). Plates were incubated at room temperature and monitored regularly. Each agar plate received a maximum of 4 individually placed single eggs. Fungi emerging from these eggs were identified using morphological and molecular phylogenetic methods. In addition they were used for studying fungal-nematode interactions in vitro. For long-term maintenance, representative fungal isolates were stored cryogenically at -140°C. Individual eggs from the second portion were directly transferred into 1.5 mL microtubes for culture-independent identification. The results of species identifications obtained from the culture-independent method were then compared to the results from the culture-dependent method. By doing so, additional evidence was gathered to prove that the fungi isolated by culturing techniques indeed colonised the nematode eggs.

Pure culture based studies

Sporulating structures were assessed after the isolates of the here studied fungal species were inoculated on corn meal agar (CMA; Fluka), oatmeal agar (OA; 30 g oatmeal, 18 g agar-agar, 1L deionised water), synthetic nutrient-poor agar (SNA; [28]), malt extract agar (MEA; Carl Roth), yeast malt agar (YMA; [29]), PDA and one third strength PDA (PDA 1/3). Strains were also inoculated on PDA, CMA and SNA supplemented with sterile pieces of filter paper, carnation leaf pieces, or wheat straw. Cultures were incubated for up to 12 months at room temperature, as well as at 10, 15, 20, 25, and 30°C in dark or under different light regimes including ambient lighting, or 12h /12 h cycles of light/darkness or black light/darkness. Growth rates at various temperatures were determined by inoculating cylindrical agar plugs, 4–5 mm diam, excised from the margin of PDA cultures onto PDA (Difco). Colour changes of fungal structures formed in culture were checked in 3% watery solution of potassium hydroxide (KOH). Colour codes used in the description were determined according to Kornerup and Wanscher [30].

Pathogenicity test and Koch’s postulates

The pathogenicity of the here studied fungal species was tested in vitro against cysts and eggs of H. filipjevi multiplied on wheat plants grown in steamed substrates in the greenhouse. Cysts were extracted and surface-sterilised for the experiments as described above. Three independently isolated strains of the fungus were sub-cultured on PDA+ and incubated for 3 months in 10 replicates. Ten surface-sterilised healthy cysts were then placed on top of each of the colonies. Plates were incubated at room temperature and cysts were monitored at regular intervals for fungal infection. Similar experiments were also done with surface sterilized eggs obtained from healthy cysts. To ensure that there is no contamination, eggs were individually placed on PDA and incubated at ambient temperature. Under a laminar flow hood, eggs not showing any contamination after 2 d of incubation, were individually placed on top or at the edge of 2-month-old PDA+, PDA1/3+ and SNA+ cultures. Plates were incubated at room temperature and eggs were monitored daily.

The process of fungal colonisation of eggs of H. filipjevi was also studied in modified slide culture experiments [31]. Single microsclerotia formed by the studied fungus (described below) were placed as inoculum in the centre of agar blocks (15×15×2 mm), and up to 20 nematode eggs were placed in their vicinity. Inoculated agar blocks were covered with sterile cover slips and slides were incubated in moist glass chambers at room temperature. Developing structures were monitored and microscopically photographed regularly.

Light and scanning electron microscopy

Nematode and fungal structures were examined and photographed with a Zeiss Axioskop 2 plus compound microscope and an Olympus SZX 12 stereo microscope equipped with a Jenoptik ProgRes® digital camera supported with CapturePro 2.8 software (Jenoptik, Jena, Germany). Eggs and fungal structures were mounted in water. Cysts were photographed in a square cavity dish in water. All microscopic specimens were studied using Differential Interference Contrast (DIC) optics. Measurements are given as minimum–maximum × minimum–maximum with arithmetical means placed in brackets, followed by the number of measurements (n).

For SEM, fungal structures of interest were either picked directly from the surface of colonies or collected after dissolving a small piece of agar in an agarose dissolving buffer (Zymo Research Corp., Irvine, California, USA). Structures were washed with SDW and placed on non-conductive double-sided adhesive tape on aluminium stubs. Samples were photographed using a FEI Quanta 250 scanning electron microscope (Hillsboro, Oregon, USA) at 12.5 kV in low vacuum. Images were adjusted in brightness and contrast using Adobe Photoshop software CS 5.1.

DNA extraction

Fungal mycelium was obtained from PDA and transferred to 1.5 mL microtubes. Genomic DNA was extracted with a CTAB-based method [32]. Cells were disrupted by grinding using sterile micro-pestles and then lysed in 800 μl CTAB buffer at 65°C for 1 h and 300 rpm. Removal of proteins and precipitation was achieved in two steps by adding 600 μl chloroform and 350 μl isopropanol. Polar fractions were retrieved through centrifugation. DNA pellets were washed twice with 70% ethanol, re-suspended in molecular grade water or elution buffer, and stored at -20°C.

Infected single eggs were transferred to 1.5 mL Eppendorf microtubes containing 5 μl SDW. Approximately 40 mg sterile silica sand and four 1 mm sterile steal beads were added. The samples were incubated in a laminar flow to evaporate the remaining water. Each sample was then frozen in liquid nitrogen and then disrupted in a tissue lyser (Qiagen TissueLyser LT, Hilden, Germany) at 50 Hz for 2 min. Freezing and disruption steps were repeated three times. DNA was then extracted and purified with the Qiagen DNeasy Plant Mini kit following the manufacturer’s instructions.

PCR amplification and sequencing

Two domains of the nuclear rDNA gene cluster including the internal transcribed spacers with the 5.8S rDNA gene (ITS) and the 5’ end of the nuclear large subunit ribosomal RNA gene (LSU) were amplified with primers V9G [33] and LR5 [34] and sequenced with primers ITS1F [35], ITS4 [36], LR0R [37], and LR5. Three partial protein-encoding genes were amplified and sequenced including the RNA polymerase II largest subunit 1 (rpb 1), actin (act), and β-tubulin (ß-tub) genes. The primers cRPB1af and RPB1cr [38] were used for amplification and sequencing of rpb1; Tact1f and Tact2r [39] were used for act, and T1 and T22 for ß-tub. For sequencing ß-tub, primer T222 and T224 [40] were used as additional internal sequence primers.

All PCR reactions were performed in a final volume of 50 μl containing 1 μl of template DNA and 49 μl of PCR master mix including 5 μl of 10× TrueStart, (NH₄)₂SO₄ amended Taq Buffer (Thermo Scientific), 5 μl MgCl₂ (2.5 mM), 5 μl dNTPs (0.2 mM of each), 2 μl of each primer (0.4 pM μl^-1), and 1 Unit Taq DNA polymerase (TrueStart Hot Start, Thermo Scientific). The amplifications were carried out on a T-GRADIENT thermocycler (Biometra, Göttingen, Germany) with the following thermal programmes: 95°C (2 min) for initial denaturation followed by 40 cycles of denaturation at 95°C (30 s), annealing at 52.5°C (ITS, LSU), 54°C (rpb1), 55.5°C (β-tub), 58.5°C (act) (40 s), extension at 72°C for 100 s (ITS and LSU), 80 s (ß-tub), and 60 s (rpb1 and act), and a final extension at 72°C (10 min). PCR products were purified using the DNA Clean & Concentrator^TM-5 kit (Zymo Research Corp., Irvine, California, USA) according to the manufacturer’s instructions. The cycle sequencing products were run on an ABI 3730XL sequencing machine (Eurofins Genomics GmbH, Germany). Obtained sequences were assembled, edited and trimmed with Sequencher 5.4.1 (Gene Codes Corporation, Ann Arbor, Michigan, USA) and deposited in GenBank under accession numbers KY607531–KY607585 and KY684180–KY684193.

Alignment and phylogenetic reconstruction

Newly generated and already published sequences were used in phylogenetic analyses (Table 1). The latter were selected according to BLASTn searches (http://blast.ncbi.nlm.nih.gov/Blast.cgi) [41] that used the former as queries. Representatives of Bionectriaceae and Nectriaceae were selected mainly following Hirooka et al. [42] and Jaklitsch and Voglmayr [43]. Several data sets based on various combinations of ITS, LSU, rpb1, act, ß-tub sequences were compiled, aligned and analysed separately. Sequences of the LSU or their combination with rpb1 (S1 Fig) or rpb1 and act (S2 Fig) were used for above genus level phylogenetic inferences. Species level phylogenetic inferences were based on all five generated loci.

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Table 1. Isolates and accession numbers used in the phylogenetic analyses.

https://doi.org/10.1371/journal.pone.0180032.t001

DNA sequences were aligned using the online version of Mafft v.7 [44] adopting the iterative refinement algorithms L-INS-I for rpb1-, act-, and ß-tub gene regions and Q-INS-i for LSU and ITS. The start and end of the alignments were trimmed manually in Se-Al v2.0 [45]. The alignments were deposited in TreeBASE, and are available at (http://purl.org/phylo/treebase/phylows/study/TB2:S20879). Based on these alignments, Bayesian Metropolis coupled Markov chain Monte Carlo analyses were done with MrBayes v3.2 [46, 47]. The general time-reversible model with the addition of invariant sites and a gamma distribution of rates across sites (GTR+I+G) was selected as the best fitting substitution model according to both the hierarchical likelihood ratio test (hLRT) and the Akaike Information Criterion (AIC) implemented in MrModeltest v2.2 [48]. Starting with a randomly selected tree, 1.000.000 (for the two- and five-gene-data set) 2.000.000 (for the three-gene data set) and 5.000.000 generations (for the LSU data set) were run, using flat prior distributions. Trees were sampled every 500 generations and 50% majority rule consensus trees were computed and a postieriori probabilities (pp) were estimated only from trees of the plateau, and after the split frequencies had fallen below 0.01. All other trees were discarded as “burnin”. The estimations were thus based on 1600 (two- and five-gene data set), 2300 (three-gene data set) and 7000 (LSU) trees sampled. Maximum likelihood (ML) analyses were performed using RAxML 7.2.8 [49, 50] implemented in Geneious 8.1.2 applying the general time-reversible (GTR) substitution model with gamma model of rate heterogeneity and 100 replicates of rapid bootstrapping (reported as MLB values). Neighbor joining analysis [51] was done in PAUP 4.0b10 in the batch file mode [52] applying the Kimura two-parameter model of DNA substitution [53] with a transition/transversion ratio of 2.0 to compute genetic distances. Support for internal nodes was estimated by 1000 bootstrap replicates [54] (reported as NJB values). The phylogenetic trees were visualised using FigTree v. 1.4.2 (http://tree.bio.ed.ac.uk/software/figtree).

Nomenclature

The electronic version of this article in Portable Document Format (PDF) in a work with an ISSN or ISBN will represent a published work according to the International Code of Nomenclature for algae, fungi, and plants, and hence the new names contained in the electronic publication of a PLOS article are effectively published under that Code from the electronic edition alone, so there is no longer any need to provide printed copies. In addition, new names contained in this work have been submitted to MycoBank from where they will be made available to the Global Names Index. The unique MycoBank number can be resolved and the associated information viewed through any standard web browser by appending the MycoBank number contained in this publication to the prefix http://www.mycobank.org/MB/. The online version of this work is archived and available from the following digital repositories: PubMed Central, LOCKSS.

Metabolite profiling

Fungal isolation

Among other nematode egg colonizing fungi to be described elsewhere, Ijuhya vitellina, newly described below, was encountered. It rendered nematode cysts collected from fields in Turkey reddish dotted upon microscopic examination (Fig 1A). Reddish dots inside cysts consisted of nematode eggs each containing one or few microsclerotia (Fig 1B and 1C). In some infected eggs, microsclerotial tissues were found developing inside juveniles (Fig 1D and 1E). When inoculated on agar medium (PDA), mycelium emerged from symptomatic nematode eggs and developed reddish orange, brick red or reddish brown cultures (Fig 1F). Predominantly globose or ellipsoidal, reddish microsclerotia were formed on the surface of (Fig 1G) or submerged in the medium. Microsclerotia formed in vitro resembled the structures encountered in nematode eggs. Cultures growing from individually inoculated nematode eggs or in vitro-produced microsclerotia developed slowly and reached a diameter of 2.8 cm within 3 months (Fig 1F, 1H and 1I). The sterility check revealed no fungal growth from the examined cysts.

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Fig 1. Cysts and eggs of Heterodera filipjevi naturally infected with Ijuhya vitellina, and pure cultures obtained from the infected eggs.

(A) Symptomatic, reddish dotted nematode cysts. (B, C) Nematode eggs accommodating reddish microsclerotia. (D, E) Microsclerotial tissue developing inside juveniles. (F) A six-month-old culture that developed from a single infected nematode egg. (G) Surface of colony showing reddish microsclerotia arranged in concentric rings. (H, I) Two-month-old cultures on PDA and CMA. Scale bars: A = 0.5 mm, B = 30 μm, C-E = 50 μm, F = 1 cm (also applying for H, I), G = 400 μm.

https://doi.org/10.1371/journal.pone.0180032.g001

Molecular phylogenetic studies

Phylogenetic reconstructions.

Alignment of 112 LSU sequences representing 100 taxa of Nectriaceae and Bionectriaceae was 860 sites long. That of 66 combined LSU (844 sites) and rpb1 (682 sites) sequences comprised 60 taxa, and that of 70 combined LSU (848), act (607) and rpb1 (684) sequences represented 64 taxa. One alignment of 15 strains representing 10 species of the genus Ijuhya only was based on all five gene regions sampled: act (624), ITS (936), LSU (819), rpb1 (712) ß-tub (875).

Strains of the nematode parasite were highly supported as a monophyletic species group in all analyses, and clustered within a highly supported clade together with other species of the bionectriaceous genus Ijuhya (Fig 2 and S1 and S2 Figs) including I. peristomialis, a later synonym of I. vitrea, which is the type species of Ijuhya [82]. Relatedness of Ijuhya with selected taxa of the Bionectriaceae, including Bionectria, the type genus of this family and various others was also highly supported. Phylogenetic analyses based on LSU only, LSU/rpb1 and LSU/rpb1/act suggested that Ijuhya oenanthicola, I. dentifera and I. antillana are distantly related to Ijuhya sensu stricto. Instead, they display closer phylogenetic affinities to Lasionectra and Ochronectria (Fig 2 and S1 and S2 Figs).

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Fig 2. Bayesian inference of phylogenetic relationships of selected taxa of the Bionectriaceae and Nectriaceae (Hypocreales) based on LSU sequences.

Numbers above nodes are estimates of a posteriori probabilities (≥ 0.9) / NJB and MLB values (≥70%). The topology was rooted with Aschersonia placenta, Balansia henningsiana, B. pilulaeformis, and Hypocrella nectrioides, (Hypocreales).

https://doi.org/10.1371/journal.pone.0180032.g002

All five loci were sampled for the Ijuhya species available as cultures to us. Hypotheses on the intra-generic phylogenetic relationships of representatives of Ijuhya were derived using these sequences. In addition two specimens of I. paraparilis from GenBank, for which three of these five gene regions were available, were added to this data set. Based on the results of the larger phylogenies, the ‘Ijuhya’ species distantly related to the Ijuhya sensu stricto were used as outgroup here. Two highly supported subclades are suggested for the in-group of genus Ijuhya, of which one includes the type species I. peristomialis, and in addition I. chilensis, I. faveliana, I. paraparilis, and I. parilis (Fig 3). The other subclade includes I. vitellina and its closest sister species, I. corynospora (Fig 3). This is in concordance with the phylogenetic hypotheses obtained from the larger two- and three-gene data sets (S1 and S2 Figs). The two isolates of I. paraparilis originating from Japan and China, did not form a species clade and are thus unlikely conspecific.

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Fig 3. Bayesian inference of infrageneric phylogenetic relationships within Ijuhya based on act, ITS, LSU, rpb1, and ß-tub sequences.

Numbers above nodes are estimates of a posteriori probabilities (≥ 0.9) / NJB and MLB (≥ 70%). The topology was rooted with three distantly related ‘Ijuhya’ species (‘Ijuhya’ antillana, I. dentifera, and ‘Ijuhya’ oenanthicola).

https://doi.org/10.1371/journal.pone.0180032.g003

Taxonomy

Ijuhya vitellina Ashrafi, W. Maier & Schroers, sp. nov., Mycobank MB 821493

Holotype for Ijuhya vitellina (here designated) (MB 821493): Turkey, Yozgat, experimental wheat field: dried culture on SNA with carnation leaf pieces, originating from an individual egg from a cyst of Heterodera filipjevi, isolated by Samad Ashrafi, August 2013 (B 70 0016479, deposited at the herbarium of the Botanic Garden and Botanical Museum Berlin-Dahlem).

Ex-holotype strain: DSM 104494, deposited in the open collection of the Leibniz-Institut DSMZ- Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH). Ex-holotype sequences: act: KY607563; ITS: KY607535; LSU: KY607549; rpb1: KY607576; tub2: KY684184.

Additional material examined, from the same location: DSM 104495 (dried culture, B 70 0016480), GenBank accession number: act: KY607564; ITS: KY607536; LSU: KY607550; rpb1: KY607577; tub2: KY684185. For other material studied, see Table 1.

Etymology: From Latin vitellus meaning egg yolk, referring to the colour and shape of microsclerotia formed by the species in nematode eggs.

Naturally infected eggs typically accommodating one, occasionally two globose to subglobose, fulvous (brownish-yellow, reddish yellow, yolk-coloured) multicellular microsclerotia with a textura angularis appearance, similar to microsclerotia formed in culture.

Colonies slow in growth; at 20°C on PDA, 6.5–9 mm diam. after 7 d, 13–16 mm after 14 d; optimum temperature for growth around 25°C, 9–12 mm (7 d), 16–19 mm (14 d); at 30°C 7–8 mm (7 d), 10–13 mm (14 d); no growth observed at 35°C. Colony reverse on OA after 21 d reddish orange (7A7), brick red to burnt Sienna (7B–D7–8) to dark brown (7F7–8) in central parts of colonies; on CMA and SNA covered with carnation leaf pieces Sahara (6C5), brick red or burnt Sienna (7B–D7–8); on up to 12 months old PDA caramel brown to brownish orange (6C6–8), cognac brown (6E7), brownish orange, brick red, copper red (7C–E7–8), or dark brown (7F6). Colony surface of similar pigmentation as reverse, granular because of solitary, gregarious or clustered microsclerotia formed on the surface of or submerged in the agar media, often arranged in concentric rings. Aerial mycelium on PDA within 3 wk not observed or sparsely to abundantly produced in sectors, white, felty to wet-cottony, on SNA and SNA with carnation leaf pieces absent or present as occasionally formed solitary, erect, typically apically coiling hyphae. Conidiophores and conidia not observed. Microsclerotia typically ellipsoidal to cylindrically oblong, sometimes globose, orange to brownish orange or brick red, not changing colour in KOH, on 6 wk-old OA 23–51 × 26–66 (36 × 44) μm (n = 52), on 6 wk-old PDA 25–46 × 32–58 (35 × 43) μm (n = 32), on ca. 12 months-old PDA cultures 26–58 × 34–77 (42 × 52) μm (n = 63). Cells of microsclerotia angular, forming a textura angularis in surface and subsurface view, variable in size, on 21 d old OA colonies 3.5–7.0 × 5.0–9.0 (5.0 × 7.0) (n = 41), on 21 d old PDA colonies 3.1–6.0 × 4.0–8.0 (5.0 × 6.0) μm (n = 50), on 12 month-old PDA colonies 3.0–7.0 × 4.5–11.0 (5.0 × 7.0) μm (n = 74) μm; walls first hyaline, later orange to brownish orange, 1.0–2.0 μm (n = 70). Microsclerotia developing from intercalary cells of hyphae or terminally at side branches, solitary or moniliform, first chlamydospore or dictyochlamydospore-like (Fig 4A–4D and Fig 5A–5C), cells angular, pigmented, first pale luteous, then brick reddish (Fig 4K–4N), filled with small guttules. Dictyochlamydospore-like structures may enlarge through coiling or expand to form globose to ellipsoidal microsclerotia (Fig 4E and 4F and Fig 5D–5F) formed solitary, in chains (Fig 4J and Fig 5G), or clusters. Globose and ellipsoidal microsclerotia developing in culture reminiscing microsclerotial structures of the fungus encountered in field-collected nematode eggs. Culturing of single microsclerotia directly extracted from nematode eggs, or retrieved from one-year old wheat straw cultures resulted in fungal growth (Fig 4O). Teleomorph not observed.

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Fig 4. Light micrographs of Ijuhya vitellina, formation of microsclerotia.

(A-F) Transformation of hyphae into (A-D) chlamydospore or dictyochlamydospore-like structures, and (E, F) microsclerotia. (G-I) Coiling or coalescence of dictyochlamydospore-like structures. (J) Microsclerotia densely arranged in a chain. (K-N) Pigmentation first observed (K) in cell walls, and later (L-M) intensifying throughout microsclerotia. (O) A single microsclerotium inoculated on agar surface developing hyphae. A-I, K-N: from PDA, J: from CMA, O: from PDA 1/3. Scale bars: (A, C, E-I, K, L, O) = 30 μm; B, J = 50 μm; D = 200 μm; (M, N) = 10 μm.

https://doi.org/10.1371/journal.pone.0180032.g004

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Fig 5. SEM micrographs of microsclerotia formed by Ijuhya vitellina.

(A) Filamentous hyphae developing into multicellular structures. (B) Intercalary formed dictyochlamydospores connected by hyphae (arrowed). (C) Detail of intercalary multicellular structures of microsclerotia. (D, E) Hyphae transformed into chlamydospore-like structures and microsclerotia. (F) Terminally formed microsclerotium. (G) Moniliform arrangement of microsclerotia. (H) Detail of microsclerotia illustrating a multicellular surface that forms a textura angularis. Scale bars: A = 100 μm; (B, E-H) = 50 μm; (C, D) = 30 μm.

https://doi.org/10.1371/journal.pone.0180032.g005

In vitro parasitism on nematode eggs and Koch’s postulates

Ijuhya vitellina infected eggs of H. filipjevi in vitro. Eggs in healthy cysts placed on colonies of I. vitellina became parasitized by hyphae within four weeks (Fig 6A). Hyphal cells inside eggs enlarged and destroyed unembryonated eggs or developing juveniles (Fig 6B). Eggs were entirely occupied by compartmentalised thick-walled hyphae (Fig 6C), and guttules-filled moniliform chlamydospores (Fig 6D) that eventually developed into microsclerotia (Fig 6E–6G). Such microsclerotia were observed within 4–5 weeks after cysts or single eggs were placed on fungal colonies; they measured 26–66 x 31–75 (41 x 49) μm (n = 97) in eggs of 3-month-old infected cysts.

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Fig 6. Light micrographs of the infection and colonisation process of Ijuhya vitellina in cysts and eggs of Heterodera filipjevi.

(A) Symptomatic cyst, reddish-dotted due to eggs containing reddish, globose microsclerotia. (B-E) Early colonisation of nematode eggs by hyphae becoming chlamydospore- and dictychlamydospore-like to develop microsclerotia inside eggs. (F, G) Dictyochlamydospore-like structures and small microsclerotia. (H) Hyphae penetrating through the eggshell by forming appressorium-like structure (arrows). (I-K) Development of the fungus inside nematodes eggs: (I) Formation of thick-walled hyphal cells, later (J-K) transforming into microsclerotia. The arrow in (J) points at the nematode stylet; in (K) at immature microsclerotium. (L) Egg with mature microsclerotium. (M-N) Near-identical cells of microsclerotium formed in (M) egg and (N) pure culture, forming a textura angulari in optical sections. Material obtained from (B-G, M) infected cysts directly placed and incubated on fungal colony, (H-L) slide cultures, (N) OA. Scale bars: A = 300 μm; (B-N) = 30 μm.

https://doi.org/10.1371/journal.pone.0180032.g006

Slide culture based observations revealed that individual eggs were infected within two weeks by hyphae that emerged from microsclerotia used as inoculum. Infection started with individual hyphae or appressorium-like structures that penetrated the eggshell and cuticle of developing juveniles (Fig 6H). Following penetration, similar infection processes and structures as described above, including swollen hyphal cells, thick-walled and multicellular structures filled with guttules, and subglobose or ellipsoidal microsclerotia were observed (Fig 6I–6L). Microsclerotia developing inside artificially infected eggs (Fig 6M and 6N) appearing textura angularis (cf. Fig 5H for details) were indistinguishable from those encountered in field-collected cysts (cf. Fig 1B and 1C).

Metabolite profiling

Structure determination of chaetoglobosins.

HRMS analysis of compounds 1 and 2 revealed the molecular formulae C₃₂H₃₆N₂O₅ and C₃₄H₃₈N₂O₆, respectively. Comprehensive analysis of the 1D and 2D NMR data of 1 and 2 indicated that compound 1 is chaetoglobosin A [16, 83] while compound 2 is its 19-O-acetylchaetoglobosin A [84] (Fig 7). The detailed description of the structure elucidation is included in the supporting information (S1 Text and S2 Text and S6 Fig).

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Fig 7.

Structures of chaetoglobosin A (1) and 19-O-acetylchaetoglobosin A (2).

https://doi.org/10.1371/journal.pone.0180032.g007

Phylogenetic analyses and systematic implications

Ijuhya vitellina is inferred as a new species on the basis of comparative morphological and molecular phylogenetic evidences. Phylogenetically, and supported by DNA sequences of five gene regions, the fungus occupies a distinct and highly supported monophyletic species clade nesting in the Ijuhya core group of the Bionectriaceae. Classification of the new species in genus Ijuhya is, however, purely based on phylogenetic evidence. While all other Ijuhya species are teleomorphically typified and largely characterized by morphological characters of ascomata and ascospores [82, 87, 88], the teleomorph of I. vitellina is unknown. Nematode associated life-style has never been described for Ijuhya species before. Other members of the genus have so far been found on plant substrata. Ijuhya vitellina differs most clearly from other Ijuhya species by the formation of brightly coloured, orange to reddish microsclerotia that have not been described for any of the other Ijuhya species. Further phylogenetic analyses on the basis of alternative taxon selections and additional data are thus required to confirm monophyly of I. vitellina with Ijuhya or resolve I. vitellina outside Ijuhya sensu stricto rendering the description of an additional genus necessary. Ijuhya vitellina, based on our taxon sampling, is most closely related to I. corynospora described from dead leaves of Phormium tenax in New Zealand (Fig 2). However, also in this case no correlating characters supporting this sister group relationship can be found. Neither were chlamydospores or microsclerotia described for I. corynospora sensu stricto [89] nor did it parasitise nematode eggs in our in vitro experiments. Also I. vitellina does form chaetoglobosins, while no such metabolites were encountered in I. corynospora or in any other of the here studied Ijuhya species. Phylogenetic analyses suggest that I. antillana, I. dentifera, and I. oenanthicola are only distantly related to Ijuhya sensu stricto and not part of the genus. Morphological characters described for I. antillana [90] and I. oenanthicola [91] conform well to the concept of the Nectria sylvana group [87], for which Lasionectria became the generic depository [82]. Specifically, fasciculate hyphal clusters formed by perithecia of I. antillana (Fig 1A, 1B and 1D in Lechat and Courtecuisse [90]) and I. oenanthicola (Plate 1, Fig. A–C in Lechat and Hairaud [91]) and occurrence of 1-septate striate ascospores are similar to those seen in Lasionectria sensu stricto. Accordingly, I. antillana and I. oenanthicola are combined into Lasionectria (Appendix).

Parasitism on nematode cysts and eggs

Our observations from field collected nematode cysts and various in vitro infection studies showed that I. vitellina parasitises cereal cyst nematodes. Several ascomyceteous fungal species have been reported to parasitise plant parasitic nematodes including cyst nematodes [2, 92–94]. Within the Bionectriaceae, to our knowledge only two species, Clonostachys rosea and Gliomastix murorum, were described as antagonists of animal- and plant parasitic nematodes [7, 95, 96].

Hyphae of I. vitellina penetrate nematode eggs either directly or by developing an appressorium-like structure. We purport that hyphae of I. vitellina may similarly enter nematode cysts or juveniles. Upon penetration the fungus forms hyphae inside eggs as is also reported for other cyst nematode parasitic fungi [2]. Penetration may apply through mechanic or chemical mechanisms [97–100]. Interactions involving chemical mechanisms are based on enzymes whose activities allow the penetration of the multilayered eggshell that in cyst nematodes mainly consists of chitin and lipids [2, 101–104]. Hyphal penetrations by I. vitellina may thus involve similar strategies to invade eggs, juveniles and cysts of H. filipjevi.

Hyphae coiling around and penetrating nematode eggs and filling the content of eggs have been described previously in various cases [105]. It is possible that hyphae directly use nutritional resources provided by nematode eggs for hyphal growth and mycelium development. The behaviour of I. vitellina inside nematode eggs differs drastically. Hyphae immediately swell and become transformed into globose or ellipsoidal dictyochlamydospores that develop into microsclerotia. Cells in these structures are angular and filled with guttules. Microsclerotia are rigid and readily resist mechanical manipulations. Additionally, guttules forming inside microsclerotia could contain lipid-like compounds for the storage of energy or may provide protection against desiccation [106]. Thus, these structures might play an important role in the survival of I. vitellina, for example during drought stress or other harsh environmental conditions. In addition, I. vitellina may not only recruit nutrients from nematode eggs for hyphal development, but inhabit these eggs inside cysts for protection and long-time survival. This is a plausible explanation as also healthy eggs of H. filipjevi can survive several years inside cysts. Empirical support for this hypothesis comes from the observation that we were (still) able to isolate I. vitellina from infected eggs after field-collected cysts were kept for several months at 4°C. The formation of microsclerotia in I. vitellina could be considered as the start and the end of the fungus’ development, at least with respect to those parts of its life cycle that were studied here. Once formed within nematode eggs, microsclerotia may remain inactive but produce newly emerging hyphae when the life-cycle of this species is newly initiated under favourable environmental conditions.

Guttules similar to those formed by I. vitellina have been suggested to serve as energy reservoirs in some other nematode parasitic fungi, e.g., the trap-forming Arthrobotrys species [107] and could also be involved in the parasitism of nematodes [105]. It was reported that guttules may contain linoleic acid as a compound responsible for nematode killing [13]. Dijksterhuis et al. [108] also suggested that microbodies, e.g., lipid organelles, present infection-related features in nematophagous fungi. However, their exact functions have not been fully elucidated. This is the first time that fungal survival structures were encountered inside nematode eggs and, accordingly, a new mode of fungus nematode interaction is described herewith.

Cysts provide a protected environment for nematode eggs where biotic and abiotic stresses are significantly reduced and eggs survive several years. Such niche might thus be a suitable environment for a nematophagous fungus where it may produce equally long-living survival structures. This situation, along with the presence of mucilaginous content of cysts, might even accelerate the fungal growth and provide optimum conditions for the fungus to colonise the entire cyst cavity and parasitise the eggs. In all cysts collected in fields, however, only a fraction of cysts carried eggs infected with microsclerotia. It is possible that I. vitellina survives in a dormant stage as microsclerotia inside cysts and that it emerges from individually parasitised eggs at favorable conditions, e.g., at times juveniles hatch from non-infected eggs within the same cyst. Microsclerotia encountered in culture have similar shapes and sizes as those I. vitellina forms in nematode eggs, either in vitro or in field collected cysts. The same applies for the cells of these microsclerotia. Accordingly this suggests that in field and in vitro encountered microsclerotia are homologous. If formed in culture they may therefore mimic the egg-parasitising habit of I. vitellina in nature. Absence of such microsclerotia in other closely related species of Ijuhya could therefore suggest that I. vitellina is the only nematode parasitising species of this genus.

Metabolite profiling

Ijuhya vitellina is reported here as a novel source of chaetoglobosin A. The vast majority of chaetoglobosins (A, B, C, D, E, F, G, and J) and their respective derivatives have mostly been isolated from the fungus Chaetomium globosum [15, 16, 109, 110]. Chaetoglobosin C is also produced by Penicillium aurantiovirens [111], and chaetoglobosin K was first extracted from Diplodia macrospora [18]. Interestingly, no such chaetoglobosins were encountered in the other, closely related Ijuhya species including I. chilensis, I. corynospora, I. faveliana, I. parilis, and I. peristomialis. Whether chaetoglobosin A and its acetyl derivative play a role in nematode egg parasitism can be inferred only with uncertainty. A temporary inhibition of mobility was observed when the two chaetoglobosins were tested in vitro against C. elegans and H. filipjevi. Chaetoglobosins affect and inhibit polymerization of actin and can degrade microfilaments [17, 112]. This might explain our observation of the effect of chaetoglobosin A and its derivative 19-O-acetylchaetoglobosin A on paralyzing the tested nematodes at 50 and 100 μg/mL. However, at higher concentrations (300 μg/mL) chaetoglobosin A was reported to have toxic effects and caused nematode mortality [26]. Thus, chaetoglobosins produced by I. vitellina may have a function in the described parasitism of nematode eggs.